The first step in the development of new drugs is the identification of molecules that bind to the target protein with relatively high affinity. This first step is usually accomplished by screening large libraries of compounds or fragments that can be used as starting points for optimization into potential drug candidates. In the case of enzyme targets, inhibition assays are usually implemented in a high throughput format in order to identify those compounds that exhibit the highest inhibition at a given concentration. In the case of non-enzyme targets, the situation is compounded by the absence of an intrinsic activity and compound binding becomes the most reliable observable for screening. In all cases (enzymes and non-enzymes) however, inhibitors need to bind to the target. Therefore, identifying those compounds that bind to the target with the highest affinity is a critical step in the identification of drug candidates. High throughput direct binding assays to arbitrary proteins have been difficult to implement. An alternative is to measure the effects of binding on specific protein properties. One such property is the structural stability of the native state of the protein. Ligands that bind to the native state of the protein will stabilize that structure; consequently, by measuring the stabilization effect of a compound on the protein target, it is possible to identify compounds that bind to the target protein. Furthermore, by measuring the magnitude of the stabilization effect, it is possible to rank the binding affinity of a library of compounds.
There are different ways to measure protein stability and each involves disrupting the protein structure through either physical or chemical means. This disruption of the protein structure is referred to as denaturation.
Temperature is one of the most widely used physical denaturants. In this scenario, a protein is subjected to increasing temperature and the corresponding changes in its structure are recorded. One of the disadvantages of temperature denaturation is that proteins typically denature at temperatures at or above 60° C. However, in most instances, the temperatures of interest are physiological (about 37° C.), room (about 25° C.) and storage (4° C.). Thus, results from temperature-based denaturation tests must be extrapolated by more than 25° C. to understand the effects at the temperatures of interest. Compounds identified by monitoring the temperature stabilization of a protein reflect the binding affinity at the denaturation temperature rather than the binding affinity at the physiological temperature. The rank order is, most of the time, different due to different temperature dependences of the binding affinity as expected from differences in the binding enthalpy.
A second way to measure protein stability is through the use of chemical denaturants, such as urea or guanidine hydrochloride. This method permits measurements to be done at any desired temperature.
The structural stability of a protein is determined by its Gibbs energy of stability, ΔG. This value, ΔG, is a function of temperature, chemical denaturants and other physical and chemical variables. Using the common example of a two state model, where a protein is either folded (i.e. native) or unfolded (i.e. denatured), the protein can transition between these two states:                NU, wherein N is the native (folded) state and                    U is the unfolded state.                        
Two different rate constants can be defined from this transitional equation. Kf is the rate of the folding reaction; while Ku is the rate of the unfolding reaction. Finally, the equilibrium constant, K, can be defined as the ratio of the unfolding rate to the folding rate, or
  K  =                    K        u                    K        f              .  Furthermore, the Gibbs energy can be expressed in terms of K, asΔG=−RT ln(K),where R is the gas constant, T is the temperature, expressed in Kelvin and ln(K) is the natural log of K. Thus, if K is greater than one, the protein unfolds at a higher rate than it folds, and its Gibbs energy is negative. Conversely, if K is less than one, the protein unfolds at a slower rate than it folds, and its Gibbs energy is positive. Also, K is equal to the ratio of the concentration of protein in the unfolded state and the concentration of protein in the folded state K=[U]/[F].
In addition, it has been observed that, for chemical denaturants, a nearly linear relationship exists between the Gibbs energy and the concentration of the denaturant. This relationship may be expressed asΔG=ΔG0−m*[denaturant],where ΔG0 is the intrinsic Gibbs energy, [denaturant] is the concentration of denaturant, and m is the multiplier, which is unique for a particular protein.
In the presence of a ligand, the Gibbs energy becomes:ΔG=ΔG0−m*[denaturant]+RT ln(1+[L]/Kd)where Kd is the binding dissociation constant of the ligand and [L] the free concentration of the ligand.
For a native/unfolded equilibrium, the fraction of protein molecules which are unfolded, or denatured, Fd, is given by:
            F      d        =          K              1        +        K              ,where K is the equilibrium constant.
This equation can be used to allow calculation of a denaturation curve. When a protein changes from its folded state to an unfolded state, certain measurable characteristics of the protein also change. One such characteristic is the fluorescence of the protein.
FIG. 1 shows a typical urea denaturation curve for an antibody. The y, or vertical, axis is a measure of the intrinsic fluorescence of the protein. The fluorescence of different dyes, usually known as protein probes, can also be used. The horizontal, or x, axis is the concentration of urea in solution with the protein. As can be seen, at a certain point, between 3M and 4M urea, the fluorescence of the protein changes dramatically, indicating that it has denatured.
While the preferred embodiment described in this application utilizes fluorescence emission (intrinsic or extrinsic) as a way to determine the degree of denaturation or unfolding of a protein, the disclosure is not limited to this technique. There are many physical observable properties and their associated instrumentation, in addition to fluorescence spectroscopy, that are sensitive to the degree of denaturation of a protein. These observable properties include, but are not limited to uv/vis spectroscopy, circular dichroism, nuclear magnetic resonance (NMR), infrared spectroscopy (IR) among others.
The generation of the data needed to produce such a graph is laborious. In one scenario, a solution containing the protein and any excipients is prepared. A sample of this solution is then subjected to fluorescent light and the emission is recorded. This is the baseline fluorescence with no chemical denaturant. In some embodiments, an amount of urea is then added to the remainder of the solution, and the light test is repeated on a portion of this modified solution. An additional amount of urea is then added to the remainder of the solution and a third light test is performed. This process is repeated for the number of desired samples. The amount of urea added each time is a function of the desired granularity of the test, and the range of urea molarities to be included. Such a method is prone to errors, as there are cumulative errors due to the constant addition of urea to the remaining solution. In this stepwise urea addition method, the process will result in the dilution of the protein and also a smaller fluorescence signal. In addition, since the solubility of urea is about 10.5M and a final 8M urea concentration is needed, the starting protein solution volume needs to be extremely small. The protein will be significantly diluted as the experiment progresses.
In another embodiment, a plurality of solutions, each with the protein, any excipients, and the proper amount of urea, is individually prepared. Each of these prepared solutions is then light tested to determine its fluorescence. While this method removes the cumulative errors associated with the previous method, it is extremely time consuming, especially for a large number of samples.
The resulting graph, such as that shown in FIG. 1, shows the stability of a particular combination of buffer, ligand and excipient conditions in the presence of a chemical denaturant. More stable combinations have a similarly shaped graph, shifted to the right. Conversely, less stable combinations have a graph shifted to the left. The presence of ligands that bind to the native state of the protein shifts the graph to the right. The magnitude of the shift is proportional to the concentration of ligand and the binding affinity of the ligand. By determining the magnitude of the shift for different potential ligands that are screened at the same concentration, it is possible to rank them in terms of their binding affinities. This is a most important goal in drug development as it provides the basis for the identification of potential drug candidates. Traditionally, full denaturation curves have been used which can be time consuming when thousands of potential ligands are screened.
It would be beneficial to create a method of identifying potential ligands which is less laborious than current processes.